Many disease processes can only be diagnosed on the basis of histologic or cytologic examination using a light microscope. For instance, while the presence of a tumor can be detected using radiological devices, the determination of whether a tumor is benign or malignant still requires a pathologist's interpretation of the appearance of the cells using light microscopy. Before reaching this stage, however, the tissue sample must first be retrieved, collected, and processed for microscopic examination. A number of techniques are available for retrieving and collecting biopsies or cell samples from a patient. It is of benefit to patients to use minimally invasive techniques for obtaining biopsies or cell samples. For example, small tissue fragments can be obtained from fine needle aspiration biopsy, or by brushing body cavity surfaces accessible through minimally invasive endoscopic techniques. Once retrieved, the cells then need to be processed for microscopy. A variety of processing techniques are known, including the CYTOSPIN technique and the THIN-PREP technique for depositing tissue fragments directly onto a microscope slide.
Another technique, commonly referred to as a cell block preparation, has several advantages over the direct deposition of tissue fragments. The cell block procedure immobilizes cells or small tissue fragments in a solid support, typically paraffin wax. Thin sections of the cell block are then cut with a microtome and the sections mounted onto a microscope slide for examination. The resulting sections from the cell block display diagnostic information in a manner that complements the direct deposition techniques. For example, the architectural arrangements of cells to each other is displayed better in sections from a cell block than in directly deposited cells on a microscope slide. Cell blocks also permit important diagnostic molecular and immunological tests to be conducted on the cell samples that would otherwise be difficult or impractical on direct preparations. In addition, cell blocks appear to preserve the cells indefinitely in a convenient manner at room temperature, thereby facilitating biomedical research.
The cell block preparation method requires that the cell fragments be “embedded” in a solid medium, most commonly paraffin wax. “Embedding” requires the following generic steps: (1) all water molecules must be removed from the cells, typically by alcohol (water is miscible with alcohol); (2) all alcohol must then be removed, as well as all fatty substances, and replaced typically by xylene (xylene is miscible with alcohol but not water); (3) the xylene must be removed and replaced with wax (wax is miscible with xylene but not with most alcohols or water); and (4) the cells in molten wax must then be manually organized and hardened on the underside of a tissue cassette so that a section of the wax block with the embedded tissue can be cut using a microtome. The first three of these steps are commonly performed by a “tissue processor,” a machine that circulates alcohol, xylene, and molten wax sequentially in a chamber containing the tissue cassette. Tissue cassettes typically serve the dual purpose of containing the cell sample during the embedding process, and for providing an attachment mechanism for holding the wax block on the microtome machine so that the cell sample subsequently embedded in wax on the undersurface of the cassette is able to be cut into thin sections.
Before “embedding” can take place, the cell sample must be manipulated to concentrate the cells and facilitate their transfer through the embedding procedure. A commonly used procedure for preparing such cell blocks is the clot technique, which is described by Yang et al. in Acta Cytologica, 42:703-706 (1998). The clot technique involves the following generic steps: (1) a cell sample is centrifuged for 10 minutes; (2) the supernatant is manually poured off, leaving a concentrated cell button; (3) fibrin and thrombin, obtained from blood bank supplies, is manually added to the cell button and incubated for 15 minutes with occasional manual swirling to trap the cell button into the clotting fibrin; (4) the clotted cell sample is removed manually from the centrifuged tube with care to avoid cell loss due to streaking the clot along the side of the tube or breaking the clot into impractically small pieces; (5) the clotted cell sample is manually transferred to lens paper, which is then folded over and placed manually into a tissue cassette; (6) the tissue cassette is then manually placed into an automated tissue processor, which then cycles alcohol, xylene and hot paraffin into the machine (as described in the preceding paragraph) and is typically set to operate overnight; (7) the following morning, the cassette is manually removed from the liquid paraffin of the tissue processor and opened; (8) the lens paper is opened, and the cell clot is scraped off the lens paper and manually placed into a tissue section mold; and (9) paraffin is gently added to the mold while manually trying to maintain the cell clot against the lowest surface of the mold that will eventually be cut. (10) The tissue cassette is then inverted over the mold to serve as a holder for the microtome machine, and hardened into the wax along with the cell clot. (11) The tissue cassette, with included wax-embedded cell fragments, is then separated from the mold. At this point, the wax block is ready to be cut with a microtome. Many of these eleven steps are common to other existing cell block production techniques.
Another popular procedure for preparing cell blocks is the collodion bag technique, which is described by Fahey and Bedrossian in Laboratory Medicine, 74(2):94-96 (1993). The collodion bag technique involves all of the eleven steps above with the exception that steps 1-4 are replaced by the following: Collodion is manually poured into a centrifuge tube to coat the tube. The cell sample obtained from the patient is then centrifuged in the coated tube. The supernatant is poured off, and the thin coating of collodion is pulled from the tube with the included concentrated cell button and embedded as in steps 5-11 above. The collodion technique provides an advantage over the clot technique by avoiding dilution of the cells with fibrin and thrombin. With the collodion technique, no waiting is required for the cell clot to form as is required of the clot technique, and cells are not susceptible to being lost as they are pulled out of the centrifuge tube. However, the collodion technique is substantially more dangerous than the clot technique due to the flammable nature of collodion and its ether solvent.
Yet another technique for preparing cell blocks is described by Diaz-Rosario and Kabawat in Cancer, 90:265-272 (2000). In this technique, the cell sample is initially filtered and the filtered sample is scraped from the filter and transferred onto lens paper which is then folded and placed into a tissue cassette and transferred to a tissue processor, followed by steps 6-11 above.
Currently available cell block preparation techniques such as the ones previously described suffer from a number of problems that make them cumbersome, costly, susceptible to contamination or mislabeling, and inefficient for showing diagnostic cells in the final microtome sections. For instance, many of the techniques require that the wax-embedded cell fragments be manually transferred to a tissue cassette required to bold the wax block onto the microtome for sectioning. This manual transfer to the tissue cassette takes a considerable amount of time, as does the step of transferring the cells from a sample tube into a tissue processor. Many of the cell fragments are wasted during the transfer and/or embedding steps. In the past, techniques attempting to improve the concentration of cells within the sectionable material and avoid cell waste have proven less than ideal, because the techniques necessarily involve dilution of the microtome section with carrier substances such that relatively few cells are present, or do not decrease cell loss in pre-embedding steps.
Current cell block techniques typically take anywhere from 8 to 16 hours to complete because the extraction of water, alcohol, fats, and xylene in the tissue processor is time consuming. While there are methods to speed up the tissue processing step (step 6 above), such as by using microwave radiation, vacuum pressure, elevated temperatures, and more rapidly diffusing chemicals, these methods suffer from their own set of problems. These techniques only modestly decrease the tissue processing time (step 6 above) but are relatively tedious to establish in the laboratory. Furthermore, these techniques do not enhance the efficiency of the other processing steps. The ability to produce same-day cell blocks would enable faster diagnoses to be rendered, with cost savings amplified throughout the health care system.
Another drawback with the cell block techniques described above are that they are also susceptible to mislabeling of a sample since the sample has to be manually moved between the sample tube, lens paper (or collodion, and other carrier substances), tissue cassette, and tissue mold. In addition to a susceptibility to complete mislabeling, cross contamination between the patients sample and cells and biomolecules (including cancer cells) of other sources is also possible with existing techniques. That is, in the fibrin clot technique, the patient's cells are mixed with pooled plasma from other patients in order to form a clot. During transfer of cells to and from the tissue cassette, contamination of cells from one case to another can occur on the forceps used to manually handle the cell buttons. Since multiple tissue cassettes are simultaneously immersed in a common bath of reagents in tissue processors, the possibility for cross contamination of cancer cells from one patient to the cell button of another sample (referred to by pathologists as “floaters”) is a well-recognized serious problem.
Yet another disadvantage of current tissue processor steps is that these steps are difficult to standardize. This is because one tissue processing machine typically circulates the embedding reagents for many laboratory samples at one time. A cell block ready to be placed on a tissue processor at 9 AM would necessarily be exposed to different conditions from a sample placed on the same processor at 9:30 AM. Emerging molecular techniques require standardized processing for optimal performance.
Since light microscopy is currently the “gold standard” for diagnosis of many diseases, advances in understanding of the molecular biology of diseases and their treatments generally requires the ability to simultaneously study the microscopic features of cells while preserving their constituent molecules for research. Simultaneous preservation of morphology and the biochemical constituents of a cell currently pose a number of problems for researchers and diagnostic pathologists (reviewed in Srinivasan M, Sedmak D, and Jewell S. “Effect of fixatives and tissue processing on the content and integrity of nucleic acids”. Am S Pathol. 2002; 161:1961-71.) Freezing tissues preserves nucleic acids and proteins (though some autolysis is inevitable during the thawing of the tissue), but frozen tissue presents obstacles to morphologic assessment. Some of the obstacles include “freezing artifacts” that distort morphology and hinder classical light microscopy diagnosis, technical difficulties in cutting thin sections of frozen tissue, inability to cut a histologic section from certain types of frozen tissues (for example tissues containing abundant fats), cumbersome and expensive storage of frozen tissue samples, impracticality of freezing very small tissue samples, and the expense of special microtomes needed to cut frozen tissues. Paraffin-embedded cell samples provide excellent morphology and biomolecules such as nucleic acids and proteins appear very stable for long term storage even at room temperature, if the biomolecules survive the embedding steps. Unfortunately, formaldehyde fixation, widely used as a fixative in production of a paraffin embedded tissue block, causes extensive damage to nucleic acids and proteins. While there are paraffin embedding techniques that do not use formaldehyde, these techniques still pose problems. The problems include the slow rate of diffusion of the embedding reagents which permits degradation of biomolecules, day to day variation in fixation conditions when a bulk embedding reagent is used over many days to embed samples, and the potential for contamination of cells from one sample into another as samples are bathed together in vats of the reagents in existing paraffin embedding techniques.
There is thus a need for a cell block preparation technique that is able to place cell fragments within a concentrated area within the cell block, without losing cells, such that a single section of the wax cell block shows a substantial proportion of all the cell fragments without dilution by fibrin or other carrier molecules. Also desirable is a technique which avoids or eliminates mislabeling of a cell block, contamination of one cell block with another cell block, and which requires less embedding reagents and time than currently available techniques, and allows standardized preservation of biomolecules for diagnostic studies and research.